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Probing Surface Adhesion Forces of Enterococcus faecalis to Medical-Grade Polymers Using Atomic Force Microscopy Annie Se´ne´chal,† Shawn D. Carrigan,‡ and Maryam Tabrizian* Department of Biomedical Engineering, McGill University, 3775 University Street, Montreal QC Canada, H3A 2B4 Received October 3, 2003. In Final Form: January 23, 2004 The aim of this study was to compare the initial adhesion forces of the uropathogen Enterococcus faecalis with the medical-grade polymers polyurethane (PU), polyamide (PA), and poly(tetrafluoroethylene) (PTFE). To quantify the cell-substrate adhesion forces, a method was developed using atomic force microscopy (AFM) in liquid that allows for the detachment of individual live cells from a polymeric surface through the application of increasing force using unmodified cantilever tips. Results show that the lateral force required to detach E. faecalis cells from a substrate differed depending on the nature of the polymeric surface: a force of 19 ( 4 nN was required to detach cells from PU, 6 ( 4 nN from PA, and 0.7 ( 0.3 nN from PTFE. Among the unfluorinated polymers (PU and PA), surface wettability was inversely proportional to the strength of adhesion. AFM images also demonstrated qualitative differences in bacterial adhesion; PU was covered by clusters of cells with few cell singlets present, whereas PA was predominantly covered by individual cells. Moreover, extracellular material could be observed on some clusters of PU-adhered cells as well as in the adjacent region surrounding cells adhered on PA. E. faecalis adhesion to the fluorinated polymer (PTFE) showed different characteristics; only a few individual cells were found, and bacteria were easily damaged, and thus detached, by the tip. This work demonstrates the utility of AFM for measurement of cell-substrate lateral adhesion forces and the contribution these forces make toward understanding the initial stages of bacterial adhesion. Further, it suggests that initial adhesion can be controlled, through appropriate biomaterial design, to prevent subsequent formation of aggregates and biofilms.
Introduction Bacterial adhesion to biomaterials remains a serious clinical problem since it can result in biofilm formation, thereby increasing the risk of infection from medical devices and implants limiting long-term success after implantation.1,2 Biofilm infections elicit complications for patients ranging from mild inflammation to subsequent surgeries and potential death, resulting in higher associated treatment costs.3,4 Since a bacterial biofilm is a complex network of cells protected by extracellular polymeric substances (EPS), antimicrobial treatment cannot efficiently reach and completely eradicate bacteria; thus, surgical intervention is necessary to remove the contaminated device or implant from the patient.5-7 Biofilm formation is a process that involves several stages and complex mechanisms.1,8-10 Bacteria are first * To whom correspondence should be addressed. Phone: (514) 398-8129. Fax: (514) 398-7461. E-mail: maryam.tabrizian@ mcgill.ca. † E-mail:
[email protected]. ‡ E-mail:
[email protected]. (1) Lamba, N. M.; Baumgartner, J. N.; Cooper, S. L. J. Biomater. Sci. Polym. Ed. 2000, 11, 1227. (2) Cadieux, P.; Watterson, J. D.; Denstedt, J.; Harbottle, R. R.; Pukas, J.; Howard, J.; Siang Gan, B.; Reid, G. Colloids Surf., B 2002, 00, 1. (3) Schachter, B. Nat. Biotechnol. 2003, 21, 361. (4) Dexter, S. J.; Camara, M.; Davies, M.; Shakesheff, K. M. Biomaterials 2003, 24, 27. (5) Anderson, G. G.; Palermo, J. J.; Schilling, J. D.; Roth, R.; Heuser, J.; Hultgren, S. J. Science 2003, 301, 105. (6) Suci, P. A.; Vrany, J. D.; Mittelman, M. W. Biomaterials 1998, 19, 327. (7) Razatos, A.; Ong, Y. L.; Sharma, M. M.; Georgiou, G. J. Biomater. Sci. Polym. Ed. 1998, 9, 1361. (8) Bowen, W. R.; Fenton, A. S.; Lovitt, R. W.; Wright, C. J. Biotechnol. Bioeng. 2002, 79, 170. (9) van der Aa, B. C.; Dufrene, Y. F. Colloids Surf., B 2002, 23, 173. (10) Woo, G. L.; Yang, M. L.; Yin, H. Q.; Jaffer, F.; Mittelman, M. W.; Santerre, J. P. J. Biomed. Mater. Res. 2002, 59, 35.
moved toward the surface by physicochemical forces, where they are reversibly attached to the surface. Irreversible binding to the surface then occurs through molecular and cellular interactions, leading to aggregation and the production of EPS, which permits stronger adhesion and cell-to-cell attachment. Finally, attached cells multiply to form microcolonies and layers. Since the initial attachment of bacterial cells on a substrate promotes biofilm development, a more thorough understanding of the mechanisms and the interactions involved in this adhesion stage could aid in delaying or preventing subsequent proliferation and colonization of the surface;1,11,12 with the aim of reducing the risk of biomaterialassociated infections, research has focused on these initial cell-surface interactions.7,8 To study the processes of bacterial adhesion, numerous experimental methods have been used that result in either the enumeration of bacteria adhered to a substrate or the estimation of bacterial adhesion strength.8,12,13 Enumeration of substrate-adhered cells has been performed by direct counting methods including light microscopy, epifluorescence microscopy, and scanning electron microscopy (SEM), and indirect counting methods such as CFU plate counting, radiolabeling, spectrophotometry, and bioluminescence analysis.8,12-14 Various forms of imaging have also been employed for the visualization of adhered bacteria using light microscopy, SEM, and confocal laser microscopy.13,14 Bacterial adhesion strength can be evaluated through the measurement of shear forces (11) Otto, K.; Elwing, H.; Hermansson, M. J. Bacteriol. 1999, 181, 5210. (12) Boyd, R. D.; Verran, J.; Jones, M. V.; Bhakoo, M. Langmuir 2002, 18, 2343. (13) Fang, H. H. P.; Chan, K. Y.; Xu, L. C. J. Microbiol. Methods 2000, 40, 89. (14) An, Y. H.; Friedman, R. J. J. Microbiol. Methods 1997, 30, 141.
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required to detach bacteria from the surface by using flow chambers or spinning disks;8,12,15 adhesion forces can also be calculated based on surface tension forces required to cause the detachment of cells by the passage of air bubbles.12,16 However, depending on the application, most of these methods suffer from limitations such as low sensitivity and precision, complexity, high cost, and lengthy procedures;13,17 further, the measurement of bacterial adhesion strength requires the estimation of a critical force value, the force required to detach all cells, increasing the number of variables and complexity of ensuing calculations.8 More recently, atomic force microscopy (AFM) demonstrated the capability of imaging bacterial cells with nanometer-scale resolution as well as the ability to probe attachment/detachment interactions with high force resolution, providing a powerful tool for the quantification of bacterial adhesion forces through the measurement of cell-cell and cell-surface adhesion forces.9,12,13,18-20 Moreover, AFM allows the analysis of biological samples in their native liquid state, precluding the need for nonphysiologic sample preparation such as chemical fixation or dehydration.9 During AFM imaging, a cantilevermounted tip is placed in contact with a surface by a specified tip-surface force; by monitoring cantilever deflection, the surface topography of the sample is acquired.8,12 The specified force is minimized when imaging bacteria to prevent damage to the sample.12,21 Current AFM methods have focused on probing cellAFM tip interactions at various cell interfaces such as the EPS that envelopes the cell9,13,22 both before and after various physical and chemical treatments.22,23 Another approach has been developed to measure tip-cell attachment/detachment forces using an AFM tip coated with a confluent monolayer of bacterial cells.8,20 However, this latter approach necessitates the immobilization of cells with glutaraldehyde and therefore does not reflect bacterial adhesion in physiological conditions.8,20 These applications give quantitative information about perpendicular forces applied between bacterial cells and the substrate, which consist of long- and short-range physicochemical forces (van der Waals forces, electrostatic interactions, chemical bonds, ionic strengths, and hydrophobic interactions) and describe the first phase of adhesion in which bacteria are brought into close contact with a biomaterial surface.24-26 Once cell-surface contact is achieved through perpendicular attraction forces, cell-surface binding is established through molecular and cellular interactions resulting from cell surface appendages (fimbriae, pili, (15) Sjollema, J.; Busscher, H. J.; Weerkamp, A. H. J. Microbiol. Methods 1989, 9, 73. (16) Gomez-Suarez, C.; Busscher, H. J.; van der Mei, H. C. Appl. Environ. Microbiol. 2001, 67, 2531. (17) Shiloh, M. U.; Ruan, J.; Nathan, C. Infect. Immun. 1997, 65, 3193. (18) Kuhlmeier, D.; Rodda, E.; Kolaric, L. O.; Furlong, D. N.; Bilitewski, U. Biosens. Bioelectron. 2002, 00, 1. (19) Beech, I. B.; Smith, J. R.; Steele, A. A.; Penegar, I.; Campbell, S. A. Colloids Surf., B 2002, 23, 231. (20) Razatos, A.; Ong, Y. L.; Sharma, M. M.; Georgiou, G. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 11059. (21) MultiMode SPM Instruction Manual, v4.31ce; Digital Instruments: 1999. (22) Camesano, T. A.; Logan, B. E. Environ. Sci. Technol. 2000, 34, 3354. (23) Camesano, T. A.; Natan, M. J.; Logan, B. E. Langmuir 2000, 16, 4563. (24) An, Y. H.; Friedman, R. J. J. Biomed. Mater. Res. 1998, 43, 338. (25) Park, J. H.; Cho, Y. W.; Kwon, I. C.; Jeong, S. Y.; Bae, Y. H. Biomaterials 2002, 23, 3991. (26) Busscher, H. J.; Poortinga, A. T.; Bos, R. Curr. Microbiol. 1998, 37, 319.
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flagella, fibrils).24,25,27 Bacterial adhesion is further strengthened by secreted extracellular substances.24,27 These surface appendages and extracellular products serve to reduce lateral mobility of adhered cells by increasing the strength of cell-substrate lateral interactions. At this stage of adhesion, the immobility of cells depends not on perpendicular interactions but on lateral interactions, which prevent environmental forces, such as shear from body fluids, from detaching bacteria.12,26 These interactions are known to be influenced by surface chemical composition and roughness and have not been studied as extensively as the perpendicular forces controlling reversible adhesion.12 Using AFM, the detachment of firmly adhered bacteria can be performed by increasing the applied force until the bending moment of the cantilever is sufficient to overcome the cell-surface adhesion force.12 This has been accomplished for the detachment of cell clusters12 but to date has not been performed on individual cells. In this work, we hypothesized that the current capabilities of this AFM technique could be expanded upon so that detachment of individual cells, and thus quantification of their adhesion strength to polymers, would be possible. This paper describes the application of the AFM to quantify the initial adhesion forces between Enterococcus faecalis, a uropathogen responsible for 80-90% of enterococcal infections,17,28 with various polymers widely used in the manufacture of medical devices and implants including polyurethane (PU), polyamide (PA), and poly(tetrafluoroethylene) (PTFE). Previous kinetics studies29 showed that PU was more prone than other polymers to E. faecalis, whereas PTFE was more resistant. Scanning electron microscopy (SEM) analyses demonstrated some differences in substrate-cell attachment depending on the nature of the polymer surface; adhesion on PU was characterized by clusters of cells and surface appendages that may strengthen adhesion, while individual cells or small groups covered PA and PTFE samples. As a means to provide further information regarding the observed differences, we investigated the potential of AFM to be used in the quantification of the adhesion forces between E. faecalis bacteria and medical-grade polymer substrates. The method developed allows for the measurement of lateral forces, contributing to the greater understanding of bacterial adhesion strength and how adhesion initiates and propagates. Materials and Methods Polymer Materials. Medical-grade polymers used in this work include polyurethane (PU) (Johnston Industrial Plastics, QC, Ca), polyamide (PA) (A. L. Hyde Company, NJ), and poly(tetrafluoroethylene) (PTFE) (Tex-O-Lon, TX). The surface dimensions of the polymeric samples were 1 cm × 1 cm. Prior to experimental studies, samples were thoroughly washed, sonicated for 10 min in distilled water and then again in 2-propanol, and finally sterilized by autoclaving for 20 min at 121 °C. Bacterial Culture. Enterococcus faecalis ATCC 29 212 were cultured at 37 °C using plate count agar (Becton Dickinson, MD) and stored at 4 °C. After inoculating 40 mL of Mueller Hinton broth (Becton Dickinson, MD) with one colony from the plate, precultures were grown overnight (18-20 h) in an orbital incubator at 120 rpm. The cell suspensions were centrifuged for (27) Dufrene, Y. F.; Boonaert, C. J. P.; Rouxhet, P. G. Colloids Surf., B 1996, 7, 113. (28) Andrews, C. S.; Denyer, S. P.; Hall, B.; Hanlon, G. W.; Lloyd, A. W. Biomaterials 2001, 22, 3225. (29) Senechal, A.; Catuogno, C. J.; Tabrizian, M. Submitted for publication.
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15 min at 3500 rpm and washed with PBS (0.01 M potassium buffer, 0.0027 M KCl, 0.137 M NaCl, pH 7.4). The final concentration was adjusted to 1.0 × 106 CFU/mL using OD546 readings (µQuant spectrophotometer, Bio-Tek Instrument). Inoculation of Polymer Samples. Prior to inoculation, polymer samples were fixed in a 2% agar mould (EM Science, NJ) in a Petri dish. Adhesion of E. faecalis on the polymer samples was then promoted by pouring a bacterial solution (106 CFU/ mL) over the samples and incubating at 37 °C in an orbital incubator at 40 rpm. At specified time intervals, samples were removed, rinsed to dislodge weakly adhered bacteria, and stored in flasks containing 2 mL of PBS for subsequent analyses. Scanning Electron Microscopy Imaging of Adhered Bacterial Cells. The distribution and morphology of the bacterial cells adhered on the polymer surfaces were studied using scanning electron microscopy (SEM) (FE SEM, S-4700, Hitachi). Prior to SEM analysis, samples in contact with the bacterial solution (1.0 × 106 CFU/mL) for a period of 3 h were prepared as described previously.29 Briefly, samples were fixed with glutaraldehyde, dehydrated with an ascending series of water-ethanol solutions followed by a series of ethanol-amyl acetate solutions, and critical point dried. Samples were then Au/Pd sputter coated (HUMMER VI, ANATECH). Scanning electron micrographs of at least three randomly selected areas of three samples of each polymer were taken. Quantification of Cell-Surface Adhesion Forces Using Atomic Force Microscopy. Samples were incubated with bacterial solution (1.0 × 106 CFU/mL) for a predetermined period of time, allowed to dry in a humid environment, and then rehydrated in deionized water for 3 h. Images of adhered bacterial cells were acquired in contact mode (NanoScope IIIa, Digital Instruments) using 200 µm long silicon nitride cantilevers having a spring constant of 0.06 N/m (NP, Veeco Instruments Ltd.); new cantilevers were used for each sample. Experiments were performed in a fluid cell filled with deionized water on at least six samples of each polymer. The AFM study was carried out as follows. Initially, the spring constant of the tip was calculated since many authors report large variability in cantilever spring constants.30 As the purpose of this study was the relative comparison of bacterial adhesion forces to different polymeric surfaces, the exact value of the spring constant was not required; the method described by the manufacturer for the determination of spring constants was considered sufficient for this application. According to this procedure, the resonance frequency of each tip was measured by centering the thermal resonance noise envelope using the cantilever tune function in the software. This frequency was then used to calculate the spring constant using
k ) Rf 3
(1)
where k is the spring constant, R is a coefficient based on tip type, and f is the measured resonant frequency. Subsequently, a force-deflection set point calibration curve was generated for each sample over a range sufficient to cover the tip force required for bacterial removal. Tip deflection in nanometers was measured for a series of deflection set points using the AFM software calibration function. The tip force exerted on the sample was then calculated as the product of the individual tip spring constant and the measured tip deflection. Lateral adhesion forces were measured by initially engaging the tip on a small area of the sample (500 nm × 500 nm) with cantilever force minimized to prevent damaging of the surface. Areas of the sample were then scanned at the minimal force (scan sizes of 20 or 40 µm) to locate individual cells. Once a bacterial cell was located, the scan size was then adjusted to frame the size of the cell and the tip scanned the cell using increasing tip force until its detachment. Figure 1 provides a schematic illustration of cell detachment using increasing AFM tip force. The detachment force was determined using the deflection set-point-applied force calibration curve. (30) Cleveland, J. P.; Manne, S.; Bocek, D.; Hansma, P. K. Rev. Sci. Instrum. 1993, 64, 403.
Figure 1. AFM tip is deflected upward in response to surface topography at low imaging force but demonstrates poor contour tracing. Increasing tip force results in improved contour tracing. Beyond the threshold of bacterial adhesion strength, the tip force is sufficient to detach cells. Table 1. Surface Properties of the Polymers before Bacterial Adhesion and AFM Results Showing the Force Required To Detach Individual Cellsa polymer
surface roughness (nm)
water contact angle
adhesion force (nN)
PU PA PTFE
8(2 20 ( 7 74 ( 15
106 ( 3 35 ( 6 117 ( 2
19 ( 4 6(4 0.7 ( 0.3
a
Results are mean ( standard deviation.
Results and Discussion The lateral force required to detach individual cells was found to be dependent on the nature of the polymer surface, with PU demonstrating the highest detachment force at 19 ( 4 nN; detachment forces for PA and PTFE were 6 ( 4 and 0.7 ( 0.3 nN, respectively. Figure 2 illustrates AFM images scanned before and after cell detachment from the different polymers, and Table 1 presents surface properties of the polymers before bacterial adhesion as well as the average calculated cell-substrate detachment force. As observed in this study, the trend of unfluorinated hydrophobic materials (PU) requiring higher detachment forces than hydrophilic materials (PA) is consistent with previous studies measuring cell-substrate perpendicular interactions,8,25,31,32 suggesting that, in general, hydrophobic surfaces may be more favorable for the initial attachment of cells to the substrate and for firmer subsequent adhesion. However, results also suggest that chemical composition also plays an important role in bacterial adhesion in addition to both hydrophobicity and nanoscale surface roughness, as the PTFE samples required the lowest cell detachment force while having the greatest roughness and lowest wettability; this lies in contrast to previous trends correlating high surface roughness to increased bacterial adhesion and biofilm formation.12,24 Previously, we investigated factors influencing bacterial adhesion to the polymers studied here, in addition to silicone (SI), high-density polyethylene (HDPE), and poly(methyl methacrylate) (PMMA)).29 Fluorescent DNAbased kinetic assays demonstrated that among those polymers, PU exhibited the most rapid initial adhesion and highest surface coverage while PTFE showed the (31) Rijnaarts, H. H. M.; Norde, W.; Lyklema, J.; Zehnder, A. J. B. Colloids Surf., B 1999, 14, 179. (32) Ong, Y. L.; Razatos, A.; Georgiou, G.; Sharma, M. M. Langmuir 1999, 15, 2719.
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Figure 2. AFM analyses of polymer samples depicting before and after images of cell detachment. Increasing surface roughness among the polymers (left to right) is evident. All images are 20 µm scale, contact mode in liquid. Circles indicate which bacteria were removed from each sample.
Figure 3. Fluorescence readings corresponding to the concentration of adhered bacteria to PU, PA, and PTFE following various incubation times (regression lines were fit from the first time point of significant adhesion). The area delimited by the dashed vertical lines represents the incubation time of polymeric samples with bacterial solution for the AFM studies. Error bars indicate standard error of the mean.
slowest adhesion kinetics and the lowest degree of surface coverage (Figure 3). Scanning electron microscopy results confirmed these differences in addition to providing visualization of polymeric surface-dependent adhesion characteristics. When considered in conjunction with these results, lateral detachment force as measured in this study provides an additional indication of an implantable polymer’s susceptibility to biofilm formation. SEM analyses clearly showed polymer-dependent differences in the morphology and distribution of adhered
cells. The PU surface was primarily covered by dense clusters of cells (Figure 4a), whereas individual cells or small groups were observed on PA (Figure 4b) and PTFE (Figure 4c) surfaces; clusters on the two latter polymers were found only in crevices. Moreover, in PU-attached clusters, surface appendages between cells as well as between cells and the substrate were found, which strengthen the cell-cell and cell-substrate interactions (see Figure 4a magnification). These protein appendages were rarely present on other polymers; on PTFE, cells
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Figure 4. SEM micrographs of E. faecalis adhered on (a) PU (bar ) 5 µm), (b) PA (bar ) 10 µm), and (c) PTFE (bar ) 5 µm); higher details shown at 25 000×.
appear to be deposited on the surface. The presence of these appendages indicates the importance of lateral interactions between the cells and a polymer surface. Cells which fail to establish adequate lateral interaction force through the presence of appendages may be more easily sheared away from the surface by flowing fluids. AFM results, having the added advantage of physiological conditions for sample preparation and image acquisition, proved consistent with SEM analysis. Singlets of cells were difficult to isolate on PU, as the surface was predominantly covered with clusters (Figure 5a), but were more easily found on polymers with lower lateral detachment forces such as PA. Confirmation of the positive lateral interaction established by bacteria on the PU surface stems from both the formation of aggregates and from the high detachment forces measured by AFM. In addition to the presence of cell clusters, measurement of single cell detachment was further complicated in some instances by the presence of an organic overlayer, likely an EPS matrix (Figure 5b). As observed by another group, this fibrillar material helps form bridges to the substrate, creating favorable conditions for cell aggregation.31 When
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the tip was engaged in a dense cluster of cells, this organic matrix periodically contaminated the tip, making subsequent high resolution difficult to achieve. Extracellular materials were noted around some individual cells adhered on PU and PA samples (Figure 5b) but rarely on PTFE, which required the lowest cell detachment force. Cell detachment experiments on PTFE samples were difficult to perform due to the limited presence of individual cells and to their weak surface adhesion. The imaging protocol required location of bacteria at 20-40 µm scan sizes before measuring detachment force using a scan size of 3-4 µm (resulting tip speed 3-4 µm‚s-1). Figure 6 illustrates a bacterial cell partially detached from PA surface. However, bacteria on PTFE frequently detached during the location phase of the protocol (resulting tip speed 40-80 µm‚s-1). Low adhesion force between E. faecalis and PTFE, likely due to the presence of fluorine in this polymer,10,33-35 necessitated performing detachment experiments on PTFE at 20-40 µm scan sizes. The impact of tip speed on the detachment force has not been examined in this study but could provide useful data in future parametric studies of lateral interaction of bacteria with substrates. To compensate for difficulties associated with tip adhesion to EPS matrix and weakly adhered bacteria, the method developed employed incubation of polymeric samples in bacterial solution for at least 4.5 h based on demonstrated time plateaus after 3 h of incubation; an incubation time between 4.5 and 5 h guaranteed that bacterial cells were firmly attached to the surface and could be easily located (Figure 3). Similar to the approach used by Yao’s group, who confirmed the biological viability of bacteria treated in their method, samples were then dried in a humid environment followed by rehydration with deionized water for 3 h.36 Initial attempts to image incubated polymeric samples without the dehydration and rehydration steps proved unsuccessful. Though the dehydration-rehydration cycle employed in this method does not replicate in vivo bacterial adhesion conditions, data obtained in this manner provide significant insights about the strength of bacterial adhesion to medical-grade polymers. To quantify the force required to detach one bacterial cell from a polymeric surface, the relationship between the applied force and the deflection set point had to be independently established for each sample analyzed. Figure 7 shows a typical force-deflection set-point calibration curve from which cell-substrate detachment forces were calculated based on linearly regressed fits. Once the applied force-deflection set-point curve was obtained, polymeric samples incubated with bacterial solution were imaged in order to find individual cells for detachment. The lateral forces demonstrated in this study are in agreement with other studies which showed that lateral forces are lower than the perpendicular adhesion forces discussed previously.8,12,26,37 To date, no comparison has been made between single-cell lateral detachment using AFM, as has been measured in this study, and alternate detachment methods such as the spinning disk and flow chamber techniques. However, the ability of AFM imaging to simultaneously locate, image, and detach bacterium (33) Woo, G. L.; Mittelman, M. W.; Santerre, J. P. Biomaterials 2000, 21, 1235. (34) Trafny, E. A.; Kowalska, K.; Grzybowski, J. J. Biomed. Mater. Res. 1998, 41, 593. (35) Grapski, J. A.; Cooper, S. L. Biomaterials 2001, 22, 2239. (36) Yao, X.; Walter, J.; Burke, S.; Stewart, S.; Jericho, M. H.; Pink, D.; Hunter, R.; Beveridge, T. J. Colloids Surf., B 2002, 23, 213. (37) Chang, K. C.; Hammer, D. A. Langmuir 1996, 12, 2271.
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Figure 5. AFM images of (a) bacterial clusters adhered on PU surface and (b) bacterial cells surrounded by extracellular substances (indicated by the arrows) on PA surface. Images are 20-µm scale, contact mode in liquid.
Figure 6. AFM image of a bacterial cell detachment from PA surface scanned from top to bottom. The image captures the upper region of the cell prior to detachment (scan size: 4 µm, contact mode in liquid).
provides unique information about adhesion strength during the initial stages of bacterial adhesion. These measured lateral detachment forces may provide more pertinent cellular adhesion information than perpendicular adhesion studies for the development of cleaning protocols for medical devices and implants.8,12 Finally, extensions of this method could promote further characterization of cell-surface adhesion, such as real-time imaging of bacterial adhesion and aggregation in physiologic conditions, and the subsequent impact of antibiotics on adhesion strength. Conclusions This study has demonstrated quantification of the initial lateral adhesion forces of individual cells of E. faecalis with medical-grade polymers including polyurethane (PU), polyamide (PA), and poly(tetrafluoroethylene) (PTFE) using unmodified AFM cantilever tips in a liquid environment. The lowest bacterial adhesion strength was found for PTFE. AFM images also qualitatively confirmed poor bacterial adhesion on PTFE in noting that no extracellular
Figure 7. Plot of the tip applied force on a PU sample as a function of the deflection set point. The regressed fit is used to calculate cell detachment force.
matrix or cell clusters were observed. Collectively, results suggest that hydrophobic surfaces may foster initial cellsubstrate contact as well as firmer subsequent adhesion, creating the potential for the formation of aggregates through surface appendages, though the chemical composition of the surface likely has a stronger influence on the overall susceptibility to biofilm formation. Lateral detachment forces measured using this method help to characterize medical-grade polymers and their resistance to microbial adhesion and subsequent biofilm formation. This greater understanding may provide insight for the standardization of adequate cleaning and sterilization protocols required for reusable medical devices. Acknowledgment. This research was made possible through funding by NSERC, MRST, FQRNT, CBB, and the French Embassy in Canada. The authors acknowledge Dr. F. Cuisinier and Dr. C. Gergely for their valuable advice in AFM analysis, K. Douglas for assistance in manuscript preparation, S. K. Sears and J. Mui for their technical expertise in SEM, and Dr. S. Prakash for access to equipment. LA035847Y
